Mouse Rif1 is a key regulator of the replication-timing programme in...

Mouse Rif1 is a key regulator of thereplication-timing programme in mammalian cells

Daniela Cornacchia1, Vishnu Dileep2,6,Jean-Pierre Quivy3,4,6, Rossana Foti1,Federico Tili1, Rachel Santarella-Mellwig5,Claude Antony5, Genevieve Almouzni3,4,David M Gilbert2 and Sara BC Buonomo1,*1EMBL Mouse Biology Unit, Monterotondo, Rome, Italy, 2Department ofBiological Science, Florida State University, Tallahassee, FL, USA,3Institut Curie, Centre de recherche, Paris, France, 4Centre National dela Recherche Scientifique (CNRS), Unite Mixte de Recherche UMR218,Laboratory of Nuclear Dynamics and Genome Plasticity, Paris, Franceand 5EMBL Meyerhof Str., Heidelberg, Germany

The eukaryotic genome is replicated according to a

specific spatio-temporal programme. However, little is

known about both its molecular control and biological

significance. Here, we identify mouse Rif1 as a key player in

the regulation of DNA replication timing. We show that Rif1

deficiency in primary cells results in an unprecedented

global alteration of the temporal order of replication. This

effect takes place already in the first S-phase after Rif1

deletion and is neither accompanied by alterations in the

transcriptional landscape nor by major changes in the

biochemical identity of constitutive heterochromatin. In

addition, Rif1 deficiency leads to both defective G1/S transi-

tion and chromatin re-organization after DNA replication.

Together, these data offer a novel insight into the global

regulation and biological significance of the replication-

timing programme in mammalian cells.

The EMBO Journal (2012) 31, 3678–3690. doi:10.1038/

emboj.2012.214; Published online 31 July 2012Subject Categories: genome stability & dynamicsKeywords: chromatin organization; DNA replication; genome

stability; replication timing; Rif1

Introduction

Replication of the mammalian genome is organized in both

space and time. Large segments of the genome, called replica-

tion domains, are coordinately replicated through the nearly

synchronous firing of clusters of replication origins. Replication

domains are visualized as foci and their position in the nucleus

as well as temporal order of activation are inherited throughout

cell cycles (Jackson and Pombo, 1998; Ma et al, 1998;

Dimitrova and Gilbert, 1999b; Sadoni et al, 2004). The

molecular and genetic mechanisms underlying such highly

orchestrated coordination and inheritance processes are still

largely unknown. Transcriptional activity, epigenetic marks

and tri-dimensional (3D) chromatin organization have all

been proposed to play a role in defining the identity of

replication domains and their order of activation (Gilbert and

Gasser, 2006; Gondor and Ohlsson, 2009; Hiratani et al, 2009).

Yet, the relationships between these inter-dependent processes

are poorly understood. A particularly intriguing relationship

is between replication timing and transcription, because

of the strong correlation between early replication and

active gene expression. A long-standing question has been

whether transcriptional activity and replication timing

are inter-dependently regulated. In some cases, changes in

transcriptional activity, for example, induced by develop-

mental transitions, have been shown to correlate with

changes in replication timing (Hiratani et al, 2008) but recent

genome-wide techniques have revealed that their relationship

is not direct (Hiratani et al, 2009). Moreover, interfering with

the activity of chromatin modifying enzymes can affect

replication timing, but the effect is generally of modest extent

and local (Aparicio et al, 2004; Li et al, 2005; Wu et al, 2006;

Jorgensen et al, 2007; Goren et al, 2008; Yokochi et al, 2009).

To date, no global determinants of the spatio-temporal

organization of mammalian DNA replication have been

identified, since all the mutations analysed influence either

few loci or repetitive sequences such as rDNA or pericentric

regions. The most dramatic rearrangements of replication

timing thus far reported take place during embryonic stem

cells (ESCs) differentiation (Hiratani et al, 2008).

The strongest correlation of chromatin properties to repli-

cation timing so far observed is to 3D chromatin organiza-

tion. Indeed, both chromatin interactions and subnuclear

position of different replication domains display a high

degree of correspondence with their timing of replication

(Gilbert et al, 2010; Ryba et al, 2010; Yaffe et al, 2010). After

metaphase, chromatin is highly mobile in the nucleus. During

G1, upon re-assembly of the nuclear envelope and lamina,

chromatin is anchored and assumes defined positions. This

moment coincides with the identification and segregation

of the replication domains whose origins fire at different

times and is therefore called ‘timing decision point’ (TDP)

(Dimitrova and Gilbert, 1999a; Leonhardt et al, 2000; Chubb

et al, 2002). One plausible theory about the mechanism of

timing control proposes the existence of a protein or protein

complex that is rate limiting for origin firing as recently

identified in budding yeast (Mantiero et al, 2011). In this

scheme, at the beginning of S-phase, origins with the highest

probability to fire would be the ones with the highest affinity/

accessibility for the limiting factor(s). Only upon release of

the limiting factor(s) from the early domains, origins with

lower affinity could then fire (Rhind, 2006). In line with the

idea of regulated access to limiting initiation factors being a

key mechanism of replication-timing regulation, it was

recently shown that the clustering of origins in subnuclear

domains mediated by Fkh1 and 2 is indeed essential for the

origin-firing programme in budding yeast (Knott et al, 2012).

*Corresponding author. EMBL Mouse Biology Unit Monterotondo,Campus Buzzati Traverso, Via Ramarini 32, Monterotondo (RM) 00015,Italy. Tel.: þ 39 06 90091348; Fax: þ 39 06 90091406;E-mail: [email protected] authors contributed equally to this work

Received: 1 February 2012; accepted: 13 July 2012; published online:31 July 2012

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3678 The EMBO Journal VOL 31 | NO 18 | 2012 &2012 European Molecular Biology Organization

The anchorage of chromosome domains to the nuclear

periphery during G1 could be a means to spatially segregate

different genomic compartments either with different protein

content and/or to spatially define the accessibility of a limiting

factor required for origin firing to a given replication domain.

Thus, by regulating interaction with the limiting factor(s),

chromatin anchoring could control the temporal order of

origin activation.

The biological significance of spatial compartmentalization

of DNA replication also remains unclear. It may be that this

organization reflects the necessity of re-assembling different

types of chromatin structures in the wake of the replication

fork, creating a temporal and local concentration of

specific factors for each chromatin type (Gilbert, 2001). For

example, temporal segregation of replication of pericentric

heterochromatin (pHC) to mid S-phase could serve the

purpose of guaranteeing the availability/concentration of

specialized chromatin remodelers and histone chaperones,

especially required for assisting fork progression through

highly compacted chromatin and for re-establishing the

appropriate epigenetic features (Quivy et al, 2008; Loyola

et al, 2009; Maison et al, 2010).

In this work, we identify Rif1 as the first key component of

the molecular machinery that determines the replication-tim-

ing programme in mammalian cells. Rif1 was originally iden-

tified in budding yeast as a Rap1 interacting protein (Hardy

et al, 1992). As for Rap1, Rif1 deletion causes telomere hyper-

elongation, suggesting that Rif1 is a telomerase negative

regulator. The mechanism by which Rif1 participates in

telomere homoeostasis remains unclear. However, it was

recently published that budding yeast Rif1 regulates telomere

replication timing by temporally restricting telomerase access

(Gallardo et al, 2011; Lian et al, 2011). In fission yeast, Rif1

does not bind Rap1, but still interacts with telomeres and

participates in their length regulation (Kanoh and Ishikawa,

2001). In mammalian cells, others and we identified Rif1 as a

double-strand break (DSBs) response factor (Silverman et al,

2004; Xu and Blackburn, 2004). In addition, we found that Rif1

had a very important role during fork re-start downstream of

ATR (Buonomo et al, 2009; Xu et al, 2010). We therefore

attributed the effects of Rif1 deletion on S-phase progression

to its activity during fork re-start (Buonomo et al, 2009). Both

during DSBs response and upon fork stalling, localization of

Rif1 at the sites of DNA damage required 53BP1 (Silverman

et al, 2004; Buonomo et al, 2009). However, Rif1 is an essential

gene (Buonomo et al, 2009), while 53BP1 knockout mice

are viable (Ward et al, 2003), prompting our investigation

of non-DNA-damage related, 53BP1-independent functions of

Rif1. Our work presented here shows that the most rapid and

widespread effect of deleting Rif1 is a genome-wide alteration

of the temporal order of origin firing. We also describe the

consequences of Rif1 deficiency on chromatin organization

and cell-cycle progression. Through studying this key

regulator, we can therefore provide an insight into the links

between replication-timing control, chromatin organization

and genome stability.

Results

Rif1 shows a dynamic S-phase-specific pattern

To gain molecular insight into Rif1 function during S-phase,

we have generated a knockin mouse replacing the starting

Rif1 ATG codon with the coding sequence of the FLAG–HA2

tag, resulting in the Rif1FH allele (Supplementary Figure S1A

and B). The functionality of this allele was confirmed by the

birth of Mendelian numbers of Rif1FH/FH mice (Supple

mentary Figure S1C) and their normal health status.

Immunofluorescence using an anti-Rif1 antibody in hetero-

zygous Rif1FH/þ mouse embryonic fibroblasts (MEFs) con-

firmed that the tagged protein fully recapitulated the

dynamics of the untagged Rif1 (Supplementary Figure S1D).

We therefore employed this endogenously epitope-tagged

Rif1 protein to study in detail Rif1 localization during cell

cycle by immunofluorescence. We performed our immuno-

fluorescence studies in cells that were pre-extracted with a

buffer containing 0.5% Triton X-100. This allowed us to

concentrate on the fraction of the protein that is either

chromatin-bound or present in the insoluble nuclear fraction.

During G1 as well as G2, apart from a variable number of

unclassified brighter foci, Rif1 was distributed diffusely

through the nucleus (Figure 1A and B). S-phase can be

divided in six different stages, each characterized by a

specific spatial EdU pattern (Supplementary Figure S2)

(Dimitrova and Berezney, 2002; Quivy et al, 2004). S1–3,

S4–5 and S6 represent early, mid and late S-phase, respec-

tively. In S1/2, Rif1 was present throughout the nucleus,

although its distribution was not homogeneous and it never

co-localized with the replication fork (Figure 2A, S1/2) or

with the replicative helicase MCM3 (Figure 1A, S1). In S3, the

latter part of early S and prior to the appearance of the

replication forks within pHC, Rif1 was specifically enriched

at chromocenters (Figure 2A), where it co-localized with

MCM3 (Figure 1A, S3). As previously described (Quivy

et al, 2004), replication of pHC occurs at the outer edges of

the chromocenters and once replicated, the DNA moves to

the interior of the chromocenter. During the initial steps of

pHC replication (S4), Rif1 localization was restricted to the

innermost of this structure, which is yet to be replicated,

appearing always closely juxtaposed to the EdU but never

overlapping with it (Figure 2B). In S5, Rif1 no longer co-

localized with the fully replicated chromocenters (Figure 2A

and B). In S6, Rif1 signal had again become more diffuse.

These data show that Rif1 displays a highly dynamic and

specific S-phase behaviour. Interestingly, Rif1 never co-loca-

lizes with the replication fork, but on the contrary, clearly

precedes it, at least at pHC during mid S-phase.

Rif1 is required for the regulation of replication timing

Rif1’s localization ahead of the replication fork, marking

domains whose origins have yet to be activated, prompted

us to investigate spatio-temporal replication patterns in Rif1

null cells. To this end, we have taken advantage of the Rif1

conditional allele (Rif1F) (Buonomo et al, 2009) to induce

acute Rif1 deletion in logarithmically growing early passage

Rif1F/F pMEFs (Supplementary Figure S3A). Upon CRE infec-

tion of Rif1F/F pMEFs, cells were pulse-labelled with EdU

and the six replication patterns typically observed in mouse

cells were identified. A low but reproducible percentage of

Rif1� /� cells displayed an aberrant EdU pattern that we

classified as mixed S2–S4 (Figure 3A). A similar observation

was reported in Suv39h1/2 double-null MEFs (Wu et al,

2006), where it was shown that the appearance of this

aberrant pattern coincided with advanced replication timing

of the major satellites. We next performed genome-wide

Mouse Rif1 controls replication timingD Cornacchia et al

3679&2012 European Molecular Biology Organization The EMBO Journal VOL 31 | NO 18 | 2012

profiling (Hiratani et al, 2008) of replication timing in Rif1þ /þ

and Rif1� /� pMEFs. Several cell cycles following CRE-

mediated deletion of Rif1, the replication-timing programme

was dramatically re-organized (Figure 3B and C), to an extent

substantially greater than that seen during ESCs differentia-

tion (Figure 3C) (Hiratani et al, 2008), the most dramatic

re-organization observed to date. The correlation between

Rif1þ /þ and Rif1� /� pMEFs was reduced to an average of

0.63, yet the correlation of replicates between two

independent lines was 0.90 (Supplementary Figure S3B and

C), demonstrating that the disrupted pattern upon Rif1 loss

was reproducible. Rif1 deletion caused both late-to-early

(LtoE) and early-to-late (EtoL) switches of replication timing

(Figure 3D and E), with 40% more domains switching from

EtoL than LtoE timing (Figure 3C). The distribution of

replication-timing values in Rif1 null cells clustered near

the middle of S-phase (Figure 3F), but due to the nature of

our genome-wide analysis, we cannot distinguish whether

these regions are replicating in mid-S, replicating randomly

throughout S-phase, or whether the two homologues are

replicating at specific but asynchronous times.

Interestingly, one striking effect of Rif1 deficiency on

replication-domain structure was their fragmentation in size

(Figure 3G). ESCs have the smallest replication domains

observed among all cell types that have been analysed thus

far, which consolidate into larger domains during differentia-

tion (Hiratani et al, 2008; Ryba et al, 2010). Rif1 deletion in

pMEFs induced fragmentation of replication-domain sizes

into a total number even higher than in ESCs (1789

domains in ESCs versus 1425 in Rif1þ /þ and 1956 in

Rif1� /�pMEFs).

These data show that Rif1 deficiency induces dramatic

changes in the temporal replication programme, to an extent

not matched by the disruption of 3D spatial replication

patterns. However, during the analysis of the spatial organi-

zation of DNA replication by EdU staining, we observed that

there were about 50% more Rif1 null cells displaying an early

S-phase pattern (Figure 4A). This result could be interpreted

as an actual accumulation of cells in the early stages of

S-phase, reflecting a temporal delay in the early-to-mid

S-phase transition. Alternatively, in many cells the pattern

that we had identified as early S could result from a loss of

spatial organization as well as a fragmentation of the replica-

tion domains. The early S-phase pattern is indeed character-

ized by a diffuse localization and by the presence of

numerous small replication foci. To distinguish between

these two possibilities, we quantified the distribution of

Rif1� /� cells among different S-phase stages by flow cyto-

metry, relying on DNA content, rather than visual inspection

(Figure 4B) to assess whether the cells were delayed in early

S-phase. We subdivided S-phase into three equal fractions

according to DNA content and determined the percentage of

total 50-bromo-20-deoxyuridine (BrdU)-positive S-phase cells

found in each fraction for wild-type versus Rif1� /� pMEFs.

This analysis revealed comparable progression through

S-phase for cells of both genotypes. We can therefore con-

clude that the visual accumulation of cells with a diffuse

early-S-like EdU pattern in the Rif1� /� cells results from

spatially and temporally disorganized replication, potentially

due to fragmentation of replication domains.

Surprisingly, we found that Rif1 deletion had no major

effect on the cell’s transcriptome (Supplementary Figure

S3D), suggesting that the effect of Rif1 deletion on replication

timing is neither mediated by nor causing gross changes in

the transcriptional landscape. However, our transcriptome

analysis has been performed on the population of cycling

Rif1� /� cells. Therefore, we cannot formally exclude that a

small number of cell-cycle specific genes could be deregu-

lated as a consequence of Rif1 deletion. The effect of Rif1

DAPIH3S10ph EdU

S6

S6

- G

2S

6 -

G2

G2

M

B Rif1FH

Mcm3 DAPIEdUA

Ear

ly G

1G

1La

te G

1S

1S

3Rif1FH Rif1FH/Mcm3

Figure 1 Rif1 localization during G1 and G2. (A) Confocal micro-scope images of cells in different stages of G1 and early S identifiedby the intensity and distribution of MCM3 signal and EdU pattern.During G1, Rif1FH (anti-HA, green) shows a diffuse nuclear staining(DAPI, blue). It co-localizes with MCM3 (red) only during the latterpart of early S (S3). EdU is shown in Cyan. Scale bar: 10mm.(B) Cells in late S-phase and different G2/M stages were identifiedby staining for phosphorylated Ser10 histone H3 (H3S10ph, red).Rif1FH (anti-HA, green) returns to its diffuse localization during G2(DAPI, blue). EdU is shown in Cyan. Scale bar: 10mm.



deficiency on the mammalian replication-timing profile that

we report here is the strongest observed to date, indicating

that Rif1 plays a key role in the control of the spatio-temporal

organization of the replication origin-firing programme.

Rif1 deficiency disrupts repackaging of newly replicated

chromatin

It has been speculated that the spatio-temporal organization

of DNA replication could facilitate the repackaging of differ-

ent types of chromatin after fork passage. Thus, one possible

consequence of replication-timing deregulation could be dis-

organized chromatin assembly (Quivy et al, 2008; Maison

et al, 2010). In agreement with this prediction, one of the

most striking phenotypes associated with Rif1 deficiency

is the presence of what we have defined ‘fluffy’ newly

replicated chromocenters. In Rif1 null cells, the EdU-

labelled DNA around the chromocenters appeared highly

disorganized (Figure 5A) in comparison to wild-type controls,

suggesting a defective pHC repackaging/assembly. However,

none of the known post-translational histone modifications

associated with heterochromatin analysed showed abnormal

localization on newly replicated pHC by immunofluores-

cence, or overall levels by western blotting (Supplementary

Figure S4A and B). We also analysed the general organization

of newly replicated DNA by examining its accessibility to

Micrococcal nuclease (MNase). For the same extent of total

DNA digestion (visualized by ethidium bromide staining),

a stronger BrdU signal in Rif1� /� pMEFs is detected

(Figure 5B; Supplementary Figure S4C), suggesting increased

accessibility of newly replicated DNA in Rif1� /� pMEFs. In

summary, Rif1 deletion causes both an alteration of the

replication-timing programme and the disorganization of

chromatin re-assembly after the passage of the replication

fork.

Replication-timing deregulation does not affect inter-

origin distances or fork progression

The profound effect of Rif1 deletion on replication timing

offered the opportunity to investigate how alteration of this

programme is related to DNA replication dynamics, namely

the frequency of origin firing and fork progression speed.

Upon CRE infection, we labelled Rif1-deficient and control

pMEFs with consecutive pulses of IdU followed by CldU to

visualize individual replication tracks by DNA Dynamic

Molecular Combing (Michalet et al, 1997). Inter-origin

distances were not significantly altered (Figure 6A) while

replication fork speeds showed a minor increase in Rif1 null

cells (Rif1þ /þ 1.98 kb/min and Rif1� /� 2.16 kb/min),

mostly resulting from a reduction in the abundance of slower

replicating forks (Figure 6B and C). Moreover, we could not

detect any difference in the number of collapsed or uni-

directional forks, nor, by flow cytometry, in the amount of

BrdU incorporated per cell (data not shown). In DT40 cells, it

has been shown that Rif1 knockout causes an increase in the

number of uni-directional replication forks (Xu et al, 2010).

However differences both in cell type and system used could

account for this discrepancy. Our data indicate that the

correct establishment of the replication-timing programme

is independent from and has no major impact on fork

progression and frequency of origin firing.

Replication-timing deregulation is a primary

consequence of Rif1 deficiency

In order to determine whether replication-timing deregulation

was upstream or downstream of the chromatin-repackaging

defect, we examined replication timing in the first S-phase

following Rif1 deletion. Rif1F/F pMEFs were arrested in

G0, deletion was induced (Supplementary Figure S4D) and

then cells were released synchronously into S-phase.

B

S3

S4

S5

A

S1/2

S3

S4

S5

EdU DAPI EdU / DAPI/

S6

Rif1FH Rif1FH/ DAPI Rif1FH/ EdU

Figure 2 Dynamic Rif1 localization during S-phase. (A) Confocal microscope images of fluorescently stained newly replicated DNA (EdU, red),immunofluorescence for Rif1FH (anti-HA, green) and DAPI (blue) in Rif1FH/FH MEFs. Scale bar: 10mm. (B) Rif1 precedes replication atchromocenters. Magnifications of insets highlighted from (A). Scale bar: 2mm.



A B

D

F G

DAPIEdU Merge

Rif

1–/–

C % Change

LtoE EtoL TOT

Control MEF versus Rif1+/+

Rif1–/– versus Rif1+/+

0.22 0.04 0.26

6.68 9.46 16.15

ESCs versus NPC*3.12 1.92 5.04

Dom

ain

size

(M

b)

0

1

2

3

4

5

6

7

Rif1+/+

Early Rif1–/–

Early ESCEarly

Rif1+/+

Late Rif1–/–

Late ESCLate

*

*

RT

log2

(ea

rly/la

te)

Rif1+/+Rif1–/– Rif1+/+Rif1–/–

0

–2

2

–4

4

Rif1+/+ Rif1–/– WT female MEF WT male MEF

Chromosome 3 (Mb)

145 150 155

RT

log2

(ea

rly/la

te)

0

–1

–2

2

1


Chromosome 17 (Mb)54 56 58 60 62 64 66

0

–1

–2

2

1

E

RT

log2

(ea

rly/la

te)

0

–1

–2

2

1


Chromosome 3 (Mb)

88 90 92 94 96 98 100

0

–1

–2

2

1


Chromosome 5 (Mb)

120 125 130

Rif1–/–

Myoblast

Rif1+/+

Male MEFNPCESC (46C)ESC (D3)iPSCESC (TT2)

Rif1

–/–

Myo

blas

t

Rif1

+/+

Mal

e M

EF

NP

CE

SC

(46

C)

ES

C (

D3)

iPS

CE

SC

(T

T2)

Figure 3 Rif1 deficiency deregulates the replication-timing programme. (A) EdU staining (green) in Rif1� /� pMEFs reveals a mixed S2–S4 patternas compared with wild-type cells (Figure 2, S2). A pan-nuclear EdU signal excluding nucleoli typical of S2 co-exists with the characteristic S4 pattern,identified by the typical EdU rings around the chromocenters. DAPI (blue). Scale bar: 10mm. (B) Hierarchical clustering and correlation matrix heatmap comparing replication-timing data (each an average of two bio-replicates) from pMEF Rif1þ /þ and Rif1� /� to different cell types (previouslypublished ESC, neural progenitor cells-NPC, Myoblasts, induced pluripotent Stem cells-iPSC) along with published wild-type MEFand two replicatesof pMEF Rif1þ /þ and Rif1� /� . (C) Table summarizing the percentages of changes comparing previously published wild-type MEFs with Rif1þ /þ

pMEFs and the latter with littermate Rif1� /� pMEFs. *As a reference, the % changes during ESCs differentiation into NPCs from Hiratani et al(2008) is shown. The percentage of changes is calculated as the number of probes on the array that change by a factor of more than 1 versus the totalnumber of probes. (D) Exemplary regions (shaded) in chromosome 17 (left) and 3 (right) whose replication-timing switches from LtoE in Rif1� /�

(red) compared with Rif1þ /þ (black) pMEFs, shown by Loess smoothed replication-timing profiles. Profiles for the same region from male andfemale published wild-type MEFs are shown in green. (E) Exemplary regions (shaded) in chromosome 3 (left) and 5 (right) whose replication-timingswitches from EtoL, depicted as in (D). (F) Distribution of replication-timing values in Rif1� /� versus Rif1þ /þ cells. Two independent lines areshown for each genotype. (G) Comparison of early (blue) and late (red) domain size distribution between Rif1þ /þ , Rif1� /� and ESCs (*ESCs fromHiratani et al, 2008).



Genome-wide analysis revealed that replication timing was

substantially deregulated already in the first S-phase after

Rif1 deletion, albeit to a lesser extent than after several cell

cycles (correlation between Rif1þ /þ and Rif1� /� 0.75,

Figure 7A–C). This result is consistent with Rif1 playing a

direct role in replication-timing determination. Interestingly,

during the first cell cycle after the induction of Rif1 defi-

ciency, the ratio between the percentages of EtoL and LtoE

switching regions is nearly equal (Figure 7A), unlike after

multiple cell cycles in the absence of Rif1, when a bias

towards EtoL emerges (1.4, Figure 3C). Hence deregulation

of the replication-timing programme is a primary effect of

Rif1 deficiency and affects both early and late replicating

domains equally.

Rif1 deficiency affects entry into the first S-phase upon

deletion

CRE-mediated deletion of Rif1 in logarithmically growing

Rif1F/F pMEFs reduced the percentage of cells actively in-

corporating BrdU by about half (Figure 8A; Supplementary

Figure S5A). This could be an indirect consequence of DNA

damage progressively accumulated during multiple cell cy-

cles in the absence of Rif1. Alternatively, entry into S-phase

could be impeded by a more direct effect of Rif1 loss. In order

to distinguish between these possibilities, we have analysed

S-phase progression of Rif1F/F pMEFs released in a synchro-

nous cell cycle after inducing the CRE-mediated deletion in

G0. As in the case of deletion in logarithmically growing cells,

the total number of cells incorporating BrdU was reduced by

about half (Figure 8B). This was not due to a defective release

from G0 to G1, as revealed by the comparable kinetics of

accumulation of cyclin D1 (Figure 8C). The appearance of a

defective BrdU incorporation already in the first cell cycle

after Rif1 deletion excludes that this phenotype originates

from accumulation of DNA damage over multiple cell cycles.

Instead, these data suggest that the defect due to Rif1

deficiency is already sensed prior to the entry into S-phase

and affects the G1/S transition (Supplementary Figure S5B

and C). To better characterize the nature of the cell-cycle

block, we analysed the behaviour of some of the crucial

regulators of the G1/S transition, such as chromatin binding

of key effectors of the pre-replication complex and p21. We

detected no difference in the kinetics of chromatin binding for

Cdc6 or Mcm3 between Rif1-proficient and -deficient cells

(Figure 8D). However, Rif1 null cells accumulated at higher

levels of p21 (Figure 8C). This is even more evident when Rif1

deletion is induced in cycling instead of contact-inhibited

cells (Supplementary Figure S5D), in agreement with the fact

that replication-timing deregulation becomes also progres-

sively more pronounced with increasing cell cycles after

deletion. p21 inhibits DNA replication both by interfering

with PCNA function (Flores-Rozas et al, 1994; Waga et al,

1994; Cazzalini et al, 2003) and by silencing Cyclin-

dependent kinases (Cdks) essential for S-phase (Sherr and

Roberts, 1999; Vogelstein et al, 2000). In agreement with a

reduced S-phase population, the amount of chromatin-bound

newly incorporated histone H4 (acetylated Lys 12 histone H4,

as an indicator for newly synthesized histones that display on

H4 the double acetylation at K5 and K12) (Figure 8D) was

also decreased (Sobel et al, 1995; Loyola et al, 2006). These

results indicate that approximately half of the Rif1 null cells

become blocked at the G1/S transition. However, half of the

cells proceed into S, fire origins at a normal frequency and

show no defect in fork progression or BrdU incorporation. At

the moment we do not know what differentiates the null cells

that arrest and the ones that don’t. Also, we don’t know if

and how the arrested cells escape the checkpoint at later

stages. However, the progressive quality of the phenotype

suggests that they do. The fact that the effect of Rif1

deficiency is already sensed prior to the beginning of DNA

replication is very interesting, suggesting the possibility that a

defective replication domain definition could be sensed by

the G1/S checkpoint.

Rif1 is associated to the nuclear matrix

Recently, budding yeast Rif1 was found to be palmitoylated

and it was shown that this modification contributes to its

association with the nuclear membrane (Park et al, 2011).

A

CRE

EV

EarlyMidLate

EarlyMidLate

52%34%

14%

75%18%

7%

53%34%

13%

56%31%

13%

0

Rif1+/+

Rif1+/+ Rif1F/F

Rif1–/–

0.5

1

1.5

2

2.5

Rat

io o

f S

-ph

ase

sub

stag

es[+

CR

E /

+EV

]

Early Mid Late

P-value = 0.28P-value = 0.18P-value = 0.08

B

Figure 4 Consequences of Rif1 deletion on spatial replication-foci distribution. (A) In the absence of Rif1, cells with early replication spatialpatterns accumulate. Upon 300 EdU pulse, cells were fixed and stained for EdU. S-phase substages were evaluated by visual inspection of thecycling population. Pie charts show the relative proportion (percentage of total S) of early, mid and late S-phase. Cells were scored blinded fortwo independent Rif1þ /þ and four Rif1F/F treated with CRE or EV pMEF clones. In all, 200 EdUþ cells were counted for each clone. Averagesare shown. P-values early Po0.0001, mid¼ Po0.0001, late Po0.01. (B) Six independent Rif1þ /þ and Rif1F/F pMEF clones treated with CRE orEVwere pulse-labelled with BrdU and the percentage of positive cells was evaluated by FACS analysis. S-phase was subdivided into three equalfractions of increasing propidium iodide content to define early, mid and late S-phase. The ratio of the percentage of total BrdU-positive cellsfound in each S-phase fraction between EV and CRE treated was plotted.



Chromatin anchoring to the nuclear membrane is a driver of

spatial organization of the chromatin in the nucleus. Since

chromatin anchoring to the nuclear periphery coincides with

the establishment of the replication-timing programme, we

determined whether mouse Rif1 was associated with the

nuclear matrix. Approximately half of total Rif1 was indeed

recovered in the lamin B-enriched fraction (Figure 9A), at

least in G1 and S-phase (Figure 9B; Supplementary Figure

S5E). Accordingly, Rif1 could also be detected in the nuclear

interior of halo preparations, displaying various distribution

patterns (Figure 9C) whose significance has yet to be defined.

To examine at a higher resolution the distribution of the

insoluble fraction of Rif1 in the cell, we performed electron

microscopy. We could detect Rif1FH signal in ultra-thin sec-

tions mostly associated to the nuclear periphery and hetero-

chromatin (Figure 9D). These data demonstrate that at least a

portion of Rif1 is constitutively associated with the nuclear

periphery. Given the fundamental role of this structure in

multiple aspects of DNA metabolism, such as DNA replica-

tion and repair, Rif1 association to the nuclear matrix could

potentially explain its involvement in both DNA repair and

replication-timing regulation. In addition, Rif1 localization is

intriguing in the light of the proposed role of 3D chromatin

organization in the definition of the temporal order of

replication domains activation.

Discussion

Although it has been recognized for a long time that

the mammalian genome replicates according to a precise

B

Rif1

–/–

A

0.06

0.08 0.

2

8

6

4

2

0B

rdU

inte

nsity

[arib

itrar

y un

its]

U MNase

WB :BrdU

EtBr

Rif1+/+ Rif1–/–

A B A B C

U MNase 0.06

0.08

0.2

0.06

0.08

0.2

0.06

0.08

0.2

0.06

0.08

0.2

0.06

0.08

0.2

Rif1

+/+

EdU DAPI Merge EdU DAPI Merge

Rif1+/+ Rif1–/–

Figure 5 Rif1 deficiency affects re-organization of newly replicated chromatin. (A) Rif1 deletion impairs post-replicative 3D re-organization ofpHC. EdU staining (green) of cells in mid S-phase shows disorganization of newly replicated pHC in Rif1� /� cells. Scale bar: 10mm. (B) BrdUpulse-labelled chromatin from two independent Rif1þ /þ and three Rif1F/F; Rosa26CreERT2/þ (Rif1� /� ) pMEFs treated with 4-hydroxytamox-ifen was digested with different dilutions of MNase. DNA was extracted and used for Southern–Western with an anti-BrdU antibody (upperpanel). The lower panel shows ethidium bromide staining of the agarose gel with the total DNA from MNase digested chromatin used for theSouthern–Western. All the pMEFs lines showed a BrdUþ population around 4%, as quantified by FACS (not shown). The right panel showsthe quantification of the BrdU immunoblot (WB) normalized for the major satellite Southern signal (SB) (Supplementary Figure S4C).U¼ enzymatic units.



spatio-temporal programme, its determinants and its biologi-

cal significance have remained elusive. The first case of a

genome-wide precocious firing of late origins in early S was

very recently reported in budding yeast (Mantiero et al, 2011).

This study showed that deregulation of replication timing

causes activation of the DNA damage checkpoint and

decreases cell viability. Here, we identify Rif1 as a key

regulator of the temporal DNA replication programme in

mammalian cells. In the absence of Rif1, both the physical

definition of replication domains as well as their temporal

order of activation is altered. The effect of Rif1 deficiency

extends throughout the genome, involving both early and late

replication domains. Currently, we do not know what

distinguishes the affected domains from the ones that are

Rif1 insensitive. We also found that Rif1 displays cell cycle-

dependent dynamic localization patterns. At least for mid

S-phase, Rif1 accumulates at sites and is cleared from

chromatin prior to its replication. It is therefore tempting to

speculate that Rif1 could be part of the mechanisms

establishing accessibility of different origin clusters for

limiting replication factors. By doing so, Rif1 would

therefore control the temporal sequence of firing of different

origin clusters. Rif1 association with the nucleoskeleton is

also compatible with this hypothesis. Intriguingly, it was

recently shown that in budding yeast, Rif1 plays a similar

role at telomeres. While telomeres are clustered and tethered

to the nuclear membrane, Rif1 imposes a time constraint (late

S-phase) for the accessibility of telomerase to the shortest

telomeres (Gallardo et al, 2011). While our work was in

preparation, it was also demonstrated that fission yeast Rif1

has a role in regulating the replication-timing programme

(Hayano et al, 2012). As we find for mammalian cells,

Schizosaccharomyces pombe rif1 deletion affects replication

timing of both early and late replication domains. This role

seems therefore to be evolutionarily conserved.

Our work also offers for the first time an insight on the

possible consequences of replication-timing deregulation on

multiple aspects of cell-cycle progression and chromatin

organization in mammalian cells. Should replication timing

serve the purpose of propagating distinct types of chromatin,

the predicted consequences of its perturbation match the

results of Rif1 deletion. We find that within the first few

cell cycles in the absence of Rif1 there are problems packa-

ging newly replicated chromatin, particularly evident at pHC.

In addition, cell-cycle progression is affected during the first

cycle after deletion. The fact that Rif1 deficiency affects the

G1/S transition is intriguing. Also, a defect preceding DNA

replication is consistent with the localization of Rif1 ahead of

the replication fork. These results are important, since they

show that deregulation of replication timing has an effect

B

0

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Speed (kb/min)

% o

f tot

al

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ed (

kb/m

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b)

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Rif1+/+

Rif1–/–

0.0 0.2 0.60.4 0.8 4.03.83.63.43.23.02.82.62.42.21.61.41.21.0 1.8 2.0

Figure 6 Rif1 deficiency does not affect the frequency of origin-firing or replication fork speed. (A) Boxplot diagram of inter-origin distancemeasured on individual DNA fibres in Rif1� /� (n¼ 38) and Rif1þ /þ cells (n¼ 53). (B) Boxplot diagram of replication fork speed measured onindividual DNA fibres in Rif1� /� (n¼ 309) and Rif1þ /þ (n¼ 392). P-value¼ 3,75 10� 5. (A, B) Bottom and top of the box indicate the upperand lower quartiles, respectively, the line in the box the median and the whiskers the 1.5 IQR of the lower and upper quartile. (C) Fork speeddistribution in Rif1� /� and Rif1þ /þ cells.



A

5.54 4.80 10.34

% Change

LtoE EtoL TOT

Rif1–/– versus Rif1+/+

B

Chromosome 3 (Mb)

145 150 155

RT

log 2

(ea

rly/la

te)

0

–1

–2

2

1

Chromosome 17 (Mb)

54 56 58 60 62 64 66

0

–1

–2

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C

RT

log 2

(ea

rly/la

te)

0

–1

–2

2

1

Chromosome 3 (Mb)

88 90 92 94 96 98 100

0

–1

–2

2

1

Chromosome 5 (Mb)

120 125 130

Rif1–/–Rif1+/+

Rif1–/–Rif1+/+

Rif1–/–Rif1+/+

Rif1–/–Rif1+/+

Figure 7 Deregulation of replication timing is an immediate consequence of Rif1 deletion. pMEFs Rif1þ /þ and Rif1F/F; Rosa26CreERT2/þ

(Rif1� /� ) were arrested in G0, 4-hydroxytamoxifen was administered in order to induce CRE-mediated deletion and the cells were releasedinto a synchronous cell cycle. (A) Table summarizing changes in replication timing in the first cell cycle after Rif1 deletion, compared withRif1þ /þ , as in Figure 3C (B) Loess smoothed replication-timing profile of the same regions (shaded) in chromosome 17 (left) and 3 (right)shown in Figure 3D whose replication-timing switches from LtoE in Rif1� /� (red) compared with Rif1þ /þ (black) pMEFs during the firstS-phase (first) upon Rif1 deletion. (C) Replication-timing profile of the same regions (shaded) in chromosome 3 (left) and 5 (right) shown inFigure 3E whose replication-timing switches from EtoL during the first S-phase (first) upon Rif1 deletion, depicted as in (B).

A B

Time (h)

CyclinD1

Tubulin

–/–

+/+

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0 13 15 17 19 21 23 h

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Tubulin

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–/–

+/+

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+/+

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+/+

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+/+

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+/+

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+/+Rif1

0 13 15 17 21 23 h

Lamin B

H4K12Ac

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15 17 19 20 21 22 23 25 27

Cdc6

Mcm3

Brd

U+ c

ells

(%)

Brd

U+ c

ells

(%)

Figure 8 The absence of Rif1 impairs entry into the first S-phase after deletion. (A) Prolonged Rif1 deletion decreases the percentage of BrdUþ

cells in an asynchronous population. A 30’ BrdU pulse was given to three independent Rif1þ /þ or Rif1F/F pMEF lines infected with a retrovirusencoding CRE recombinase (CRE). Bars show the average percentage of BrdUþ cells. (B) Rif1 deletion was induced as in Figure 7. S-phaseprogression was monitored by FACS analysis of BrdU content and the percentage of BrdUþ cells over the entire population was plotted versustime. An average of three independent pMEF lines for each genotype is presented. (C) Western blotting of Cyclin D1 and p21. Loading control:a-tubulin. h¼hours. (D) Western blot of chromatin-bound Cdc6, MCM3, acetylated Lys12 histone H4 (H4K12-Ac). Loading control: lamin B.(C, D) One representative pair of Rif1þ /þ ; Rosa26CreERT2/þ and Rif1F/F; Rosa26CreERT2/þ (Rif1� /� ), bothþ 4-hydroxytamoxifen treated areshown.



immediately prior to the initiation of DNA replication, but not

on its progression. The frequency of origin-firing and fork-

progression parameters are indeed mostly unaffected by Rif1

deficiency. We have observed that Rif1 deletion reproducibly

decreases the abundance of the slower replication forks.

However, we do not know if this is a direct consequence of

Rif1 deficiency, or, alternatively, the result of altered

chromatin status or a form of cell adaptation.

Previous studies with immortalized Rif1 null mouse and

chicken cells detected activation of the DNA replication

checkpoint, accumulation of chromatid breaks (Buonomo

et al, 2009) and problems in fork progression (Xu et al,

2010). Comparison among these different data sets suggests

that the immortalization process (perhaps loss of the G1/S

checkpoint) allows cells a certain degree of tolerance to Rif1

deficiency (our unpublished data) with the consequent

accumulation of replication stress and genomic instability

(Buonomo et al, 2009). Mutations found in the Rif1 gene in

breast cancer cell lines (Sjoblom et al, 2006; Howarth et al,

2008) have raised the possibility that deregulation of Rif1

function could be involved in cellular transformation. Our

data provide potential mechanisms, by illustrating how loss

of Rif1 function determines profound changes of the

replication-timing programme, activation of the G1/S

checkpoint and chromatin-repackaging defects. Each of

these events on its own has been shown to either identify

cancer cells (De and Michor, 2011; Ryba et al, 2012) or to

contribute directly to cell transformation (Bartkova et al,

2005; Gorgoulis et al, 2005; Halazonetis et al, 2008; Zhu

et al, 2011). In conclusion, the work we report here identifies

Rif1 as one of the first key mammalian factors discovered in

the yet unknown network that temporally controls DNA

replication and, as a consequence, genome stability.

Materials and methods

Mice generationThe Rif1F mouse allele has been described previously (Buonomoet al, 2009). For the generation of the mouse Rif1FH allele, the

second exon, containing the starting ATG, was replaced with FLAG–HA2 tag containing an ATG. The targeting vector was generatedby recombineering (Lee et al, 2001). We inserted a FRT-NeomycinR(NeoR)-FRT cassette downstream of the second exon.The first FRT site is associated to a diagnostic NdeI restriction site,the second to BamHI. The construct was subcloned into the pDTA-TKIII vector that allows double-negative selection (DTA and TKgenes), linearized with NotI and electroporated into Bruce4 C57BL/6 ES cells. ES cells colonies were screened for homologousrecombination by Southern blot, digesting the DNA with BamHIand using the R5 probe. Positive clones were further confirmed byNdeI digest and injected by standard techniques into C57BL/6Jblastocysts. Chimeras were evaluated based on tail genotyping.Chimeric founders were crossed to C57BL/6J females and themice were kept in a pure C57BL/6J background. The NeoRcassette was removed by crossing the F1 mice with a FLPe deletermouse strain (Jackson Labs). Genotyping of mice and derived cellswas performed by PCR with the following primers: f1: GCGAACCTCGGACGCCGTGG; r1: GCACCTGTAATCTCAACCACTC. PCRgenerates a 146-bp product for the wild-type allele and a 248-bpproduct for the Rif1FH allele. The Southern probes R5 (848 bp) wasgenerated by PCR from genomic DNA with the following primers:Rif1FH50fw 50-GTGTCACTACTCTCACATTT and Rif1FH50rev 50-TGTTTTTCCATTTAGAAGCCAG.

Cell manipulationpMEFs were generated from E12.5 embryos according to standardprotocols and propagated in DMEM high Glucose, 15% heat-inacti-vated fetal bovine serum (PAA), 100U/ml penicillin, 0.1mg/mlstreptomycin, 0.2mM L-glutamine, 0.1mM non-essential aminoacids and 50mM b-mercaptoethanol. pMEFs at passage 2 wereinfected as described in Buonomo et al (2009), except that weperformed 5 instead of 4 infections, spaced 12–6–6–6–12 h. After 4days of selection in Hygromycin B (Sigma H3274) at 90mg/ml cellswere processed without further passaging.Cell synchronization was performed by driving early passage

pMEFs into contact inhibition and then inducing CREERT2 transloca-tion into the nucleus by treating the cells with 600nM 4-hydro-xytamoxifen (Sigma H7904). During the last 2 days of treatment, thecells were also serum-starved.

ImmunofluorescenceCells were pulsed with EdU for 300, and then processed as describedin Buonomo et al (2009). EdU staining was performed according tothe manufacturer’s instructions (Invitrogen Click-iT kit C10350).Evaluation of cells in different S-phase substages for the different

A

Rif1

Tubulin

Histone3

LaminB

TS DSI DSII DR HSS HSR

B

Rif1

TS TIG0/G1

TS TIS

TS TIG2/M

CDAPI Merge

D

HC

NE

NE

Rif1FH

Figure 9 Rif1 is associated with the nuclear matrix. (A) Western blotting on extracts from MEFs fractionated for nuclear matrix isolation. Anti-Rif1 antibody used was 1240. TS¼ triton soluble; DS¼DNase soluble; DR¼DNase resistant. The DR fraction was further divided in HSS¼highsalt soluble and HSR¼high salt resistant. (B) Live MEFs were FACS sorted in three fractions (G1, S and G2) on the basis of their DNA contentusing Hoechst 33342. The cells collected were fractionated in triton-soluble (TS) and -insoluble fraction (TI). Western blot was performed onextracts from equal number of cells for each fraction using anti-Rif1 antibody 1240. The migration shift of Rif1 TI fraction band is probablycaused by the presence of UREA 8M (see Materials and methods). (C) Nuclear halos were prepared from Rif1FH/FH MEFs. Immunofluorescencefor Rif1FH (anti-HA, green) and DAPI (blue). Scale bar: 10mm. (D) Electron microscopy visualization of Rif1FH in triton pre-extracted MEFs. Onerepresentative ultra-thin section per each cell and magnification of insets highlighted are shown. The black dots correspond to the Nanogoldparticles associated to the secondary antibody. HC, heterochromatin; NE, nuclear envelope.



lines was performed blind. Images were acquired using a Leicaconfocal TCS SP5 microscope with a � 63 1.4 NA oil objective andrun by LAS AF Software (Leica). Contrast adjustment and croppingwere performed in Photoshop (Adobe) and ImageJ. Figures werecomposed in Illustrator (Adobe).

Preparation of halosHalos were prepared according to Guillou et al (2010). Anti-HAimmunofluorescence was performed as above.

Micrococcal nuclease treatment and Southern–WesternCells were pulsed for 1 h with 10 mM BrdU and collected. Nucleiwere prepared by dounce homogenization in lysis buffer (20mMTris–HCl pH 7.5, 4mM MgCl2, 250mM sucrose, 0.1% Triton X-100and protease inhibitors). After pelleting them 50 at 2000 g at 41C,nuclei were resuspended in lysis buffer and chromatin was isolatedover a sucrose cushion (Lysis buffer with additional 30% sucrose)at 2400 g 150 41C. The isolated chromatin was resuspended inMNase buffer (60mM KCl, 15mM NaCl, 15mM Tri–HCl pH 7.4,0.25M sucrose, 1mM CaCl2) and digested for 2min with theindicated amounts of Micrococcal nuclease (Sigma N3755). Thereaction was stopped with stop buffer (50mM EDTA, 2.5% sarcosyl,1% SDS) and proteins digested with Proteinase K 10mg/ml. Equalamount of DNAwere loaded on a 1.3% agarose gel prepared in Tris-Glycine buffer (5� : 144 g Glycine, 30 g Trisbase per liter) and usedfor Southern blotting according to standard protocols. The mem-brane was crosslinked twice at 0.15 J/cm2 and washed in 0.1MNaOH. After 1 h blocking in 3% BSA in PBS-T (Tween 0.1%), themembrane was incubated in anti-BrdU antibody for 1 h. The samemembrane was probed for Southern with the major satellite probefrom Lehnertz et al (2003).

Western blotting, chromatin fractionation and nuclear matrixisolationTotal protein extracts and western blotting were performed asdescribed in Buonomo et al (2009).Chromatin fractionation was performed by incubating cells in

CSK buffer (0.5% Triton X-100, 100mM NaCl, 3mM MgCl2, 300mMsucrose, 1mM EGTA, 10mM PIPES pH 6.8) 10’ on ice then spinning4 at 500 g. Supernatants were recovered and represent the Triton-soluble (TS) fraction. Pellets were washed twice in CSK buffer,resuspended in 8M Urea and represent the Triton-insoluble (TI)fraction.Nuclear matrix isolation was performed as in Mladenov et al

(2006).

AntibodiesAnti-HA (Covance HA.11 Clone 16B12) was used at 1:3000;Anti-BrdU (BD 347580) 1:2000 for western blotting 1:7 for FACS;Anti-Cdc6 (Cell Signaling 3387 Clone C42F7) 1:1000; Anti-p21(Santa Cruz sc-6246 1:200); Anti-CyclinD1 (Santa Cruz M-20, sc-718) 1:200; Anti-MCM3 (Santa Cruz N-19, sc-9850) 1:200 forwestern blotting, 1:50 for immunofluorescence; Anti-H3 (CellSignaling 9681) 1:1000; Anti-H3K9me3 for western blotting(Novus Biologicals 6F12H4) 1:1000; Anti-H3K9me3 (Abcam,ab8898) for immunofluorescence 1:500; Anti-HP1a for westernblotting (Upstate 07-346) 1:1000; Anti-HP1a for immunofluores-cence (Euromedex 2HP-1H5-AS) 1:500; Anti-H3K27me3 (Abcam,ab6147) 1:1000; Anti-H4K20me3 (Abcam ab9053); Anti-H4K12Ac(Cell Signaling 2591) 1:1000; Anti-Lamin B (Abcam, ab16048)1:1000; Anti-MeCP2 (Abcam, ab2828) 1:1000; Anti-phospho-Histone H3 (Ser10) (Millipore 06-570) 1:200; Anti-Rif1 (Buonomoet al, 2009) 1:8000 for western blotting and 1:3000 for immuno-fluorescence; Anti-Smc1 (Bethyl Laboratories A300-055A) 1:5000;Anti-a-Tubulin (Sigma T9026) 1:10 000; Anti-g-Tubulin (SigmaT6557) 1:5000.

FACS analysis and sortingBrdU labelling, staining and FACS analysis were performed as inBuonomo et al (2009).For FACS sorting logarithmically growing MEFs were stained with

Hoechst 33342 and sorted in the three fractions according to theirDNA content on a 5-laser ARIA SORP (BD Biosciences). Oncecollected, the cells were pelleted and the triton-soluble/insolublefractions were obtained as described.

DNA combingNeo-synthesized DNAwas labelled with an IdU pulse followed by aCldU pulse as described in Anglana et al (2003). Genomic DNAwasextracted and combed on silanized slides as in Letessier et al (2011).Immunofluorescence detection of neo-synthesized DNA andDNA was performed by successive incubations and washes as inAnglana et al (2003) with the following reagents: (1) mouse anti-bromodeoxyuridine (BrdU) (BD Biosciences) at 1:5 dilution and ratanti-BrdU (AbD Serotec) at 1:10 dilution; (2) Alexa-488-conjugatedgoat anti-mouse (Invitrogen) and Alexa-594-conjugated goat anti-rat (Invitrogen) both at 1:50 dilution; (3) mouse anti-single-stranded DNA (Chemicon) at 1:25 dilution; (4) Alexa-594-conjugated goat anti-rat (Invitrogen) and Alexa-647-conjugatedgoat anti-mouse (Invitrogen) both at 1:50 dilution. Data weregenerated with two independent Rif1þ /þ and Rif1� /� pMEFpreparations.A Leica DMR600 epifluorescence microscope equipped a

CoolSNAP HQ2 CCD camera and run by Metamorph software(Molecular Devices) was used for image acquisition with a � 40objective. Overlays of IdU/CldU/DNA signals performed inMetamorph were used to determine CldU replication-track lengthand inter-origin distances. Only CldU signal (second pulse) juxta-posed to IdU signal was used to determine replication speed.Replication tracks with interruption of the DNA signal or with alength identical to the DNA signal were not considered since theycould reflect broken fibres. Replication fork speed was calculated bydividing the length of a CldU track (in kb) with pulsing time (200).Statistical analysis was performed with a Mann–Whitney test.

Transcriptome analysisTotal RNA was isolated using the TRIzol reagent (Invitrogen)following the manufacturer’s instructions and treated with DNAse(Promega). RNA quality was checked using a bioanalyzer (Agilent2100; Agilent Technologies), and RNA quantity was measured withND-1000 Nanodrop spectrophotometer. In all, 1mg of RNA samplewas used for microarray analysis Affymetrix Mouse Gene 1.0STarray(Affymetrix). Robust multi-array average (RMA) normalization wasapplied. Normalized data were then filtered based on the Affymetrixdetection call so that only probes that had a Present call in at least oneof the arrays were retained. CEL files were then imported inGeneSpringGX 11.5 software and statistical analysis was performedto detect significantly differentially expressed genes. A fold changecutoff of 2 was applied.

Replication profilingNewly synthesized DNA was pulse-labelled with (BrdU) while thecells were in the exponential phase. Cells were then dissociated intoa single-cell suspension and nuclei were isolated. DNA was subse-quently stained with propidium idodide and cells were FACS sortedinto early and late S-phase fractions based on their increasing DNAcontent during S-phase. The DNA that was synthesized either earlyor late during S-phase was then purified by immunoprecipitation ofthe BrdU-substituted nascent DNA (BrdU-IP). For the analysis ofreplication timing in the synchronous cell cycle, cells were collectedat each time point after 2 h BrdU pulse. In this way, we ensured thatonly cells in the first S-phase after Rif1 deletion were analysed. Afterfixation and before FACS sorting, the different time points werepooled. Thus, the definition of early and late S-phase was based onthe DNA content and not on the time of collection. A series of 10DNA sites that are known to replicate at specific times were thenanalysed by PCR to verify the quality of the nascent strands, whichwas then subjected to whole-genome amplification. The amplifiedearly and late fraction were differentially labelled and hybridized toa CGH whole-genome tiling array. The replication timing of eachprobe along the genome was then calculated as the log2 enrichmentof early fraction over late fraction. Genome-wide replication-timingprofiles were constructed as described (Hiratani et al, 2008) withexception that Rif1� /� pMEF data set was not scaled to thestandard Inter-quartile range (IQR) of other data sets. This wasdue to the unusually narrow distribution of the majority of datapoints, which caused the outlining data points to go beyond scaleduring standard scaling. Since the scale of Rif1� /� data wassignificantly different from Rif1þ /þ in both replicates, this pointsto a biological cause. Two replicates of Rif1� /� and Rif1þ /þ

pMEFs were averaged and used to create the replication-timingprofiles. The replicates of Rif1þ /þ data and Rif1� /� data were



normalized separately using Limma package in R. Then, the datawere Loess smoothed using 300Kb window. After smoothing, the2–98 percentile range of Rif1� /� was scaled to 2–98 percentilerange of Rif1þ /þ . This scaled data were used for further analysis.

Electron microscopyRif1FH/FH MEFs were plated to 80% confluency on gridded MatTekdishes (MatTek, Ashland, USA). Cells were then rinsed with PBS,pre-extracted with Triton X-100 buffer pH 7.9 (0.5% Triton, 20mMHepes-KOH, 50mM NaCl, 3mM MgCl2, 300mM sucrose) for 10minat 41C. They were then rinsed with PBS and fixed with 3%paraformaldehyde and 2% sucrose in PBS for 10’ at room tempera-ture. Cells were then rinsed with 50mM NH4Cl, extracted with 0.5%saponin and incubated with the EdU reaction mix for 30’. After theincubation, they were rinsed with PBS and imaged on a ZeissAxioObserver epifluorescence microscope and S-phase nucleiwere imaged. Cells were then blocked, incubated with an anti-HAantibody (Covance, Princeton, USA) for 1 h, rinsed and then in-cubated with an anti-mouse IgG Nanogold antibody overnight.They were then further processed for electron microscopy asdescribed in Bahtz et al (2012). Thin sections (60 nm) wereimaged on formvar-coated slot grids and stained with 2% uranylacetate in 70% methanol and 1.5% lead citrate. Imaging was doneon a CM120 Phillips microscope.

Supplementary dataSupplementary data are available at The EMBO Journal Online(http://www.embojournal.org).

Acknowledgements

We gratefully acknowledge Violetta Parimbeni for mousehusbandry, Claudia Valeri for genotyping and Melanie Leuener fortechnical help. This study was technically supported by EMBLMonterotondo’s FACS and microscopy and EMBL HeidelbergGenomics Core Facility. We also acknowledge the services ofJ Rientjes from Monash University’s Gene Recombineering Facilityand Patrick Varga-Weisz (Babraham Institute, Cambridge, UK) forthe Micrococcal assay protocol and useful discussions. We greatlyappreciate for helpful discussions Donal O’Carroll (EMBLMonterotondo). We are especially grateful to Nadia Rosenthal. GAand JPQ received support from la Ligue Nationale contre le Cancer(Equipe labellisee Ligue 2010), the European Commission Networkof Excellence EpiGeneSys (HEALTH-F4-2010-257082), ERCAdvanced Grant 2009-AdG_20090506 ‘Eccentric’, ANR ‘ECenS’ANR-09-BLAN-0257-01. RF was supported by EMBL Interdiscipli-nary Postdoc (EIPOD) under Marie Curie Actions (COFUND); SCBBthanks the EpiGeneSys Network of Excellence.Author contributions: DC performed the majority of the experi-

ments. VD performed the replication-timing experiment startingfrom the FACS sorting of the cells and analysed the data. JPQperformed the DNA combing and its analysis. RF did the transcrip-tome analysis. FT helped in several experiments and performed theBrdU FACSes. RM and CA have carried on the electron microscopyanalysis. GA and DG have given scientific feedback on both theexperiments and the manuscript preparation. SCBB designedthe experiments and wrote the paper.

Conflict of interest

The authors declare that they have no conflict of interest.

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